De Novo Engineering of Solid-State Metalloproteins Using

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De novo engineering of solid-state metalloproteins using recombinant coiled-coil silk Trevor D. Rapson, Tara D Sutherland, Jeffrey S Church, Holly E Trueman, Helen Dacres, and Stephen C. Trowell ACS Biomater. Sci. Eng., Just Accepted Manuscript • Publication Date (Web): 23 Sep 2015 Downloaded from http://pubs.acs.org on October 6, 2015

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ACS Biomaterials Science & Engineering

De novo engineering of solid-state metalloproteins using recombinant coiled-coil silk Trevor D. Rapson,†* Tara D. Sutherland, †* Jeffrey S. Church,‡ Holly E. Trueman, † Helen Dacres§ and Stephen C. Trowell† †

CSIRO, Black Mountain, Canberra, ACT, 2601, Australia

‡

CSIRO, Waurn Ponds, Geelong, VIC, 3216, Australia

§

CSIRO Werribee, VIC, 3030, Australia

Abstract In order to achieve the sophisticated chemistry required for life, nature uses metal containing proteins (metalloproteins). However, despite intensive research efforts, very few of these metalloproteins have been exploited for biotechnological applications. One major limiting factor is the poor stability of these proteins when they are removed from their cellular environment. To produce stable metalloproteins, we have developed an engineering strategy that uses structural proteins which can be fabricated into a number of different solid-state materials. Here we demonstrate that a recombinant silk protein (AmelF3 – Apis mellifera Fibroin 3) binds heme and other metal macrocycles in a manner reminiscent of naturally occurring metalloproteins, whereby an amino acid coordinates directly to the metal center. Our strategy affords design at four different levels; the metal center, the organic macrocycle, the protein scaffold and the material format structure. The solid-state metalloproteins produced remained functional when stored at room temperature for over one year.

Keywords: heme, recombinant silk, biosensors, biocatalysis, industrial biotechnology

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Introduction The engineering of metalloprotein mimics is attracting increasing interest both as a route to understanding natural protein structure-function and for a range of biotechnological applications.1–3 The heme protein family is of particular interest due to the diversity of chemical functions accessible through the interaction of the iron heme metal center with a variable protein scaffold (Figure 1a).4–6 The ability to harness the functional properties of heme proteins within a solid-state material affords a number of potential practical advantages, such as stability and recyclability, for heterogeneous biocatalysts and biosensors.7

Figure 1 a Comparison between naturally occurring heme protein structures and engineering scaffolds. Cartoon structures of a common heme protein with an α-helical structure (myoglobin – pdb, 2VIK), the de novo maquette scaffolds used to produce engineered metalloproteins (pdb, 1M3W)8 and the predicted coiled-coil structure of recombinant honeybee silk (AmelF3) modelled from tropomyosin (pdb, 1C1G). b Comparison of the molecular structure of AmelF3 (Apis mellifera Fibron 3 ~ 30 kDa) and silkworm silk (heavy fibron ~ 250 kDa). The β-sheeted regions of silkworm silk are made up of repetitive regions containing i – GAGAGS ii - GAGAGY and/or AGAGVGY, iii similar to (i) except for the presence of an AAS motif. The minimum structural requirements to mimic native metalloprotein function has been investigated using de novo engineering of metal binding sites into α-helical peptides that form


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bundles in solution (Figure 1a).2,6,9 This approach has produced artificial molecules capable of oxygen-binding,10 light harvesting,11 transmembrane transport12 and catalysis.5,13–15 In this work we sought to transfer the insights gained by de novo engineering from the solution-state to solid-state materials. In particular we have focused on the use of transgenic honeybee silk (AmelF3) as a new protein scaffold for metalloprotein engineering.16 A number of recent reports have described the engineering of structural proteins to bind metals.17–1920,21 This approach has allowed new functions to be introduced into structural materials. For example, the addition of short silver binding peptides to spider silk produced a chimeric protein which could be cast into films with anti-microbial properties.22 Through covalent attachment of titanium binding peptides to silkworm silk, enhanced cellular adhesion to titanium based implants was achieved.17 Krishnaji and Kaplan developed a chimera between a uranium peptide recognition motif and spider silk, which has potential applications in the area of bioremediation.18 Although in these examples the structural proteins can be fabricated into solid-state materials, the goal was not to mimic metalloprotein function but rather produce a material capable of binding metals such as uranium for bioremediation,18 or for releasing metal ions, as in the case for antimicrobial activity.22 In contrast, here we sought to control the reactivity of the metal center via direct interaction with the protein scaffold, to carry out functions such as catalysis.23 Honeybee larvae spin silken cocoons that contribute to the mechanical and thermal stability of the hive.24 The honeybee silk proteins, unlike silkworm and spider silk, adopt a molecular structure with a large central coiled coil domain (~210 αα) with short (20-50 αα) β-sheeted regions at the N and C termini (Figure 1b).25,26 This coiled coil structure is similar to the αhelical de novo maquettes (Figure 1a) successfully used by other groups to engineer metalloprotein mimics. While natural honeybee silk is made up of four fibrous proteins,27a single fibroin of honeybee silk (Apis mellifera Fibron 3 -AmelF3) can be fabricated into 3 ACS Paragon Plus Environment


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various solid-state materials including fibres,28,29 sponges30 and films.31 Importantly, AmelF3 can be produced at high levels (>2g/liter) in recombinant systems16 making commercial production viable. The extensive natural variation found among honeybee silk homologs32 suggests that the AmelF3 fibroin amino acid sequence can be substantially varied to insert or modify metal and macrocycle binding sites without comprising the proteins ability to be fabricated into material forms. In this work we describe the use of AmelF3 as an engineering scaffold to mimic naturally evolved heme-containing proteins. Heme proteins are attractive targets for de novo engineering,5,9,33 given the diverse functions they perform using a common iron porphyrin ring. One of the principal ways that the reactivity of naturally occurring heme proteins is controlled is through direct coordination of the iron by an amino acid residue.34 The importance of direct interaction between the protein scaffold environment and metal center to metalloprotein function has become increasingly clear.23,35–37 Our immediate goal was to achieve both binding and direct coordination of heme cofactors to AmelF3. If coordination can be achieved, this would allow further tuning of reactivity through varying the identities of the coordinating residue and the metal macrocycle. Experimental section Materials All standard chemicals used were purchased from Sigma Aldrich (St Louis, MO) unless stated otherwise. Heme b, copper and cobalt protoporphyrin IX and tetrasulfonate zinc phthalocyanine were purchased from Frontier Scientific (Logan, UT). Cobyrinic acid heptamethyl ester was purchased from Sigma Aldrich. UV/Vis absorption measurements were

carried out on a SpectraMax M2 plate reader.


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Silk protein preparation Recombinant honeybee silk protein, AmelF3 was produced by fermentation in Escherichia coli as described by Weisman et al.16 Briefly, purified inclusion bodies were solubilised by addition of equal dry weight sodium dodecyl sulphate (SDS) from 1 – 10% solutions as required. The majority of the detergent was then removed by centrifugation after addition of 300 mM KCl which led to its potassium-induced precipitation. This treatment generated high-purity AmelF3 solutions containing approximately 0.3% SDS weight/volume. The AmelF3 solution was dialysed against 15% polyethylene glycol (PEG) to generate a solution containing 3.6% protein and 0.3% SDS. Metal macrocycle solution preparation Heme b (iron protoporphyrin IX), cobalt protoporphyrin IX and copper protoporphyrin IX were initially dissolved in a minimal amount of 0.1 M NaOH (~ 1 mL) and made up in 70% methanol to give a final volume of 10 mL. Other macrocycles such as cobyrinic acid heptamethyl ester and tetrasulfonate zinc phthalocyanine were dissolved directly in 70% methanol. Concentrations of ~ 0.05 mg/mL were used for all macrocycles. Metal functionalised silk material preparation AmelF3 silk protein sponges were generated as follows; AmelF3 protein solution was poured into silicone rubber molds (14 x 5 x 6 mm; RL060, ProSciTech, QLD), frozen at -20ºC overnight, and placed in a freeze-dryer (FD355DMP, FTS Systems) for 24 hours to generate sponges with typical dimensions of 12.6 x 4.5 x 5.4 mm. The silk sponges were stored in sealed plastic bags at room temperature until use. Silkworm silk sponges were a gift from Prof David Kaplan.



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Heme b (or other macrocycles) could be added to the honeybee silk or silkworm silk sponges by adding solutions of the macrocycle (in 70% methanol) drop-wise to the sponge (~ 10 mg protein). Aqueous 70% methanol was used as the solvent for AmelF3 materials as they are not soluble in 70% methanol. An alternative method of preparing silk-heme sponges was to place a freeze-dried sponge into a 10 mL solution of macrocycle and allow the sponge to absorb the solution until fully saturated. The sponge was washed 3 times with 70% aqueous methanol (10 mL per wash), air-dried and stored at room temperature. Silk films were prepared by either co-drying heme and silk protein, or by leaching heme into preformed films. Co-drying involved dissolving the equivalent of 1 mg heme b with 10 mg of recombinant honeybee silk sponges in 0.5 mL of hexafluoroisopropanol (HFIP) overnight at room temperature. The solution was aliquoted into either a cuvette or a 24-well plate, air dried at room temperature and stored until required. The dried film was soaked overnight in 70% methanol to make the film insoluble in water. To add heme b (or an other metal macrocycle) to a preformed film, the film was first prepared by dissolving a sponge in milliQ water (~ 10 mg silk in 1 mL of water) and then a drop (~ 100 uL) of the solution was air dried on a plastic surface to generate a disk film. The film was soaked in 1 mL of the macrocycle solution (0.05 mg/mL) for 48 hours followed by washing with 70% methanol to remove unbound heme. Substituting tyrosine in AmelF3 Site directed mutagenesis was carried out using Pfx50 DNA polymerase from Invitrogen (Carlsbad, CA), with overlapping primers to introduce the desired mutation (see supplementary information) according to the manufacturer’s instructions. The template plasmid was removed by Dpn1 digestion following PCR amplification. Sequencing was carried out at the Micromon sequence facility (Monash University, Australia) using T7


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primers. Try76Ala and Tyr76His substituted protein was expressed and purified as described for the unmodified silk protein. Nitric oxide binding assay Heme b-silk films were covered with 50 mM phosphate buffer (pH 7) and reduced by the addition of ~ 10 µL sodium dithionite (100 mM) in phosphate buffer (pH 7). Diethylamine 2nitrosohydrazine sodium hydrate (NONOate) was dissolved in 50 mM phosphate (pH 7) to generate 1.5 mol equivalent of dissolved NO. Aliquots of the diethylamine NONOate solution was added incrementally to a dithionite reduced film under argon gas. Changes in the UV-Vis spectrum were monitored to determine redox states and measure NO binding. The experiment was carried out in triplicate using separate films allowing standard deviations to be calculated for each point. Peroxidase assay The protocol outlined by the Worthington Biochemical Corporation at http://www.worthington-biochem.com/hpo/assay.html was adapted as follows. A solution of 1.7 mM H2O2 (prepared from a 30% solution purchase from Sigma (St Louis, MO) was mixed with a solution of 2.5 mM 4-aminoantipyrine (JT Baker (Centre Valley, PA)) and 170 mM phenol (Sigma (St Louis, MO)) in a 1:0.9 ratio. Heme b-silk sponges were added to this solution and absorbance at 510 nm was monitored over time. In this assay Fe3+ is oxidized by hydrogen peroxide and then reduced by 4-aminoantipyrine (hydrogen donor), which in turn oxidatively couples to phenol to form a pink colored quinone. Raman spectroscopy Raman spectra were obtained using a Renishaw inVia confocal microscope system with 754 nm excitation from a Renishaw HPNIR diode laser through a x50 (0.75 na) objective. Incident laser power, as measured using an Ophir Nova power meter fitted with a photodiode



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head, was 4.93 mW for the silk films and 0.66 mW for the heme b powder. Films were held on a mirrored backing. The powder was compressed into a 2 mm cavity cell. A coaxial backscatter geometry was employed. Spectra were collected over the range of 100 to 1900 cm-1 and averaged over at least 4 scans, each with an accumulation time of 20 seconds. For the films, six spectra collected from different sample areas were averaged to produce the final spectrum. Raman shifts were calibrated using the 520 cm-1 line of a silicon wafer. The spectral resolution was approximately 1 cm-1. Fourier transformed infrared spectroscopy Infrared attenuated total reflectance (ATR) spectra were obtained using a Perkin Elmer (Beaconsfield, UK) Spectrum 100 Fourier transform infrared (FTIR) spectrometer equipped with a universal micro-ATR accessory and purged with liquid nitrogen boil off. Spectra were collected at a resolution of 4 cm-1 by co-averaging a minimum of 16 interferograms. Manipulation of spectral data was carried out using Grams AI v 9.1 software (Thermo Fischer Scientific, USA). Results and Discussion Initially we tested the binding of the most commonly occurring heme cofactor, heme b, to freeze-dried honeybee silk sponges. Given the coiled coil structure of AmelF3 and the large number of positively charged amino acid residues such as lysine and arginine, we expected that heme b would bind weakly to the unmodified honeybee silk protein. However, when a honeybee silk sponge was exposed to heme b solutions, which are green, the sponge immediately turned red (Figure 2a – top panel), indicating a change in the coordination of the iron metal center within the heme group. Furthermore, the binding of heme b to honeybee silk was found to be strong; the cofactor could not be removed by washing the heme b – silk sponges with aqueous solutions, 70% methanol in water (Figure 2a – top panel), 0.1 M hydrochloric acid or 0.1 M sodium hydroxide and organic solvents such as chloroform.


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Figure 2 Production and optical properties of a solid-state biomimetic metalloprotein. (a), Evidence for specific binding and coordination of heme b within honeybee silk sponges. A solution of heme b was added to a honeybee silk sponge (top panel) and a silkworm sponge (bottom panel). The sponges were then washed with 70% methanol. (b) Heme b in 70 % methanol (top) could be diffused into a preformed film to yield a heme b containing silk film (bottom). (c) Cartoon representation of heme b binding and coordinating to AmelF3 giving rise to the color change in the heme b cofactor from green - uncoordinated to red coordinated. (d) UV-visible spectrum of a heme b-silk film compared to that of heme b in aqueous solution. A video of the addition of heme b solutions to AmelF3 and silkworm silk is provided in the supplementary materials.

In contrast, when heme b was added to sponges composed of silkworm (Bombyx mori) cocoon silk proteins, the green solution was absorbed by the sponge by the heme b solution was readily washed out with 70% methanol (Figure 2a – bottom panel, supplementary video).



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Our results clearly indicate that heme b was able to bind to AmelF3 but not to silkworm silk sponges. The majority of naturally occurring heme proteins such as the globins and cytochromes have an α-helical structure (e.g. myoglobin Figure 1a). In these proteins, the heme cofactor is held within the protein through a combination of hydrophobic packing, ionic or hydrogen bonding between the heme propionate groups and charged or polar amino acid residues such as arginine, tyrosine or serine.3,38 It is therefore likely that the coiled-coil structure of AmelF3 assists with the binding of heme b by providing a hydrophobic environment. Furthermore AmelF3 is rich in charged residues such as lysine (7.5%) or hydrogen bond donors such as serine (11.3%) which can interact with the propionate groups of heme b. In addition to the strong binding of heme b by AmelF3, the color change, from green to red, noted upon the addition of heme b to honeybee silk sponges (Figure 2a) indicates a change in the coordination of the iron metal center within the heme group. To investigate this further, we analyzed the Ultraviolet-Visible (UV-Vis) spectra of heme b diffused into preformed transparent films (Figure 2b-2d). Heme proteins have a characteristic Soret peak at ~ 400 nm, which is sensitive to changes in the coordination of the iron heme atom. In the case of iron porphyrins, such as heme b, broad Soret peaks below 400 nm indicate that the iron is coordinated to the porphyrin ring (4-coordinate) but not the protein. When the heme group is also coordinated to an amino acid in the protein the Soret band red shifts above 400 nm and sharpens, as a five-coordinate iron center is generated.39 The UV-Vis spectrum from the honeybee silk-heme b films show a sharp Soret peak at 404 nm (Figure 2d) confirming that the iron heme center is coordinated to the honeybee silk protein (AmelF3). These results indicate that AmelF3 silk materials bind and coordinate heme b in a manner resembling naturally occurring heme proteins.


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Given that we observed that heme b both binds and coordinates to AmelF3, we sought to determine the effect of adding heme b on the molecular structure of AmelF3 in the form of a film. Fourier transformed infrared spectroscopy (FTIR) allows the secondary structure of protein materials to be investigated using the amide I and III modes.40,41 The FTIR spectra of both dried films and those stabilized with 70% methanol were measured to compare films with and without heme b (Supplementary Figure S1). No conformational differences were detected due to the addition of heme b (Figure S1). As previously described for dried AmelF3 materials16 both spectra of the dried films exhibit amide I peak maxima at 1646 cm-1 which is consistent with that expected for a dominant coiled coil structure. A weaker low wavenumber shoulder was also observed, indicative of a minor amount of β-sheet structure. The amide III region exhibits peaks at both 1304 cm-1 (coiled coil) and 1235 cm-1 (β-sheet). Following stabilization with 70% methanol, similar to that reported previously,16,30 the spectrum of both the stabilized films with and without heme were found to be very similar with amide I peak maxima at 1644 and 1626 cm-1. This is consistent with increased β-sheet structure (1626 cm-1). We have previously determined by infrared spectroscopy that methanol treatment results in around 10% decrease in coiled coil conformation with a corresponding increase in β-sheet content.16 To allow further engineering options it was important to understand exactly which residues of AmelF3 where coordinating to the iron center of heme b. AmelF3 does not contain any of the coordinating residues typically found in natural heme proteins, such as histidine, cysteine or methionine (Supplementary Figure S2). Therefore, to identify the coordinating amino residue, we investigated the stoichiometry of heme binding and coordination. By co-drying heme and protein solutions, rather than diffusing heme b into preformed films, we were able



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to prepare films with varying ratios of heme b to silk protein. At low heme loadings (heme: protein molar ratios of 1:1 and 1:2), a sharp Soret peak was observed at 404 nm, indicating that the majority of the heme was coordinated (Figure 3a). As the ratio of heme b to protein was increased, the Soret peak broadened and shifted to lower wavelengths, indicating an increase in the amount of uncoordinated heme. This finding suggests that a single site within each silk monomer is responsible for heme coordination.

Figure 3 A tyrosine in honeybee silk specifically coordinates to heme b. (a) UV-Vis spectra of honeybee silk films containing varying amounts of heme b. Inset shows the change in the position of the Soret peak with differing molar ratios of heme b. (b) Raman spectra from honeybee silk (grey trace) heme b-silk (red trace) and heme b powder (green trace) excited at 785 nm. (c) UV-Vis spectra of films made from Tyr76Ala silk compared with unmodified honeybee silk films. Inset show photos of unmodified silk sponge (red) and Tyr76Ala sponge (green) following washing with 70% methanol.

Raman spectroscopy was used to measure stretching frequencies between the iron center and the coordinating ligand, which are indicative of the identity of the ligand.42 The Raman spectrum of the silk-heme film, excited at 785 nm, showed a broad peak centered at 594 cm-1 (red trace, Figure 3b). Heme proteins that have a tyrosine coordinating ligand show similar Fe-Tyr stretches.43,44 AmelF3 contains a single tyrosine residue (Tyr76) located in the core of the coiled coil region. To test whether Tyr76 is the coordinating ligand, we replaced Tyr76 12 ACS Paragon Plus Environment

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with alanine using site directed mutagenesis. When heme b was added to sponges formed from Try76Ala modified protein, the sponge did not become red (Figure 3c – inset) suggesting that no coordination change to the heme cofactor was occurring. Furthermore, the UV-Vis spectrum of transparent films generated from the Tyr76Ala substituted protein had a broad Soret peak at 395 nm indicating that the coordination observed in unmodified honeybee silk had been reversed by this single amino acid substitution (Figure 3c). Nevertheless, the uncoordinated (green) heme b could not be washed out of the sponge, indicating that the cofactor still bound to the Tyr76Ala silk, even though the iron center was not coordinated to the silk (Figure 3c – inset). This binding is most likely mediated via interactions between the propionate group of heme b and charged or polar residues of AmelF3 and the hydrophobic core of the coiled-coil domain. Although silkworm silk has over 4% tyrosine residues, none of these residues coordinates, or binds, heme b in the manner observed for AmelF3 (Figure 2). This result suggests that in addition to the presence of a coordinating residue, the molecular architecture of AmelF3 plays an important role in both the binding and coordination of heme b. To further demonstrate that the binding site could be modified, we used site directed mutagenesis to change the coordinating tyrosine residue to a histidine, the residue most commonly found in natural heme proteins.4,45 A Tyr76His modification induced a red shift of the Soret peak to 410 nm in films generated from this recombinant AmelF3 material (Supplementary Figure S3), consistent with coordination of histidine to the metal centre.39,46 Uncoordinated heme and heme-like cofactors have been widely used in biomimetic chemistry. Their lack of coordination, however, has limited their usefulness.7 Using AmelF3, as described here, we can vary the coordination of the heme center, allowing more precise control over the functional properties of the metalloproteins.



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To demonstrate the catalytic potential of solid-state metalloproteins we engineered a heterogeneous catalyst based on a heme b silk sponge. Heme peroxidases catalyze the breakdown of phenol derivatives, one of the largest groups of pollutants in industrial effluents.47–49 The catalytic mechanism involves peroxidase oxidation by hydrogen peroxide followed by reduction of the peroxidase by a substrate such as a phenol or aromatic amine (Figure 4a). We tested the catalytic activity of heme b-silk materials using the oxidative coupling of phenol with 4-aminoantipyrine to produce a pink quinone with λ max = 510 nm (Figure 4b, Supplementary Figure S4). The heme b-silk sponges readily catalyzed the coupling of phenol to 4-aminoantipyrine in aqueous media. Furthermore, the sponges were recoverable, they could be reused at least six times (Figure 4c) and remained functional if stored dry at room temperature for over one year.

Figure 4 Demonstration of catalytic activity by heme b-AmelF3 materials. (a) The catalytic mechanism of heme b-silk peroxidase activity. 4-aminoantipyrine acts as the substrate reducing the heme center. (b) Time course of phenol oxidation by heme b-AmelF film. Coupling of phenol to 4 amino-antipyrine gives a colored quinone. Inset shows the change in the solution visible spectrum over time. (c) A heme-AmelF3 sponge was transferred through a series of wells containing 4-aminoantipyrine (1.1 mM), phenol (80 mM)


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and H2O2 (0.9 mM). The rightmost well contains a solution to which the sponge has not yet been added. The second well from the right contains the sponge.

As a second example of the properties of solid-state metalloproteins we used a heme b-silk film as a nitric oxide sensor. The films were sensitive to nitric oxide at sub-micromolar concentrations and the binding was reversible (Supplementary Figure S5). In addition, the sensing property was retained in films stored dry at ambient humidity and room temperature for over a year (Figure 5). Poor stability at room temperature of many biologically derived materials is one of the factors that has prevented the transition of metalloproteins from the research laboratory to commercial applications. This work demonstrated that stable metalloprotein biosensor materials can be produced through the use of a stable protein as the engineering scaffold.

Figure 5 Heme b-silk films as a stable biosensor. The UV/Vis spectra of a reduced heme bsilk film before (solid line) and after (dashed line) the addition of nitric oxide. Left panel: freshly prepared film. Right panel: nitric oxide binding of the same film after storage for one year at ambient conditions.

We have demonstrated the ability to produced heme b-AmelF3 materials in different formats such as sponges and films. The reactivity of the iron heme center can be altered through varying the coordinating ligand from AmelF3. To broaden our engineering options, we 15 ACS Paragon Plus Environment


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extended our survey beyond heme cofactors to other metal macrocycles. Biological systems incorporate a range of non-heme metal macrocycles in proteins, such as the magnesium chlorophylls in Photosytem II50 and the cobalt corrins in Vitamin B12.51,52 Inspired by nature’s metal cofactors, synthetic chemists have produced a range of compounds such as the metallophthalocyanines which are used in dye-sensitized solar cells53,54 and photodynamic therapy.55 Varying the metal center and organic cofactor allows additional avenues of engineering to produce solid-state materials with different functional properties (Figure 6a).

Figure 6 Engineering options for the development of solid-state metalloproteins

(a) Schematic representation of the four different levels of engineering control available to produce solid-state materials for a diverse range of applications. (b) Examples of solid-state materials developed: AmelF3 films with i – Copper protoporphryin IX and ii – Cobalt protoporphyrin IX. AmelF3 sponges with iii – Cobyrinic acid heptamethyl ester and iv – Tetrasulfonate zinc phthalocyanine (molecular structures shown in Figure S6). AmelF3 sponges with heme b v unmodified, vi – Y76A substitution. In all cases solid-state materials were thoroughly washed with 70% methanol to remove unbound macrocycle. vii – AmelF3 fibres (woven), viii – AmelF3 transparent films (no metal cofactors added).

We therefore sought to determine if AmelF3 materials could be engineered to incorporate other cofactors besides iron heme. We were readily able to replace the iron in heme with copper or cobalt (Figure 6b), which can alter functional properties. For example, cobalt16 ACS Paragon Plus Environment

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substituted hemes have been used as selective nitric oxide sensors.56 In addition to hemes, other organic macrocycles, such as corrin and phthalocyanine rings (Supplementary Figure S6), bind to AmelF3 materials (Figure 6b). The use of the versatile family of metallophthalocyanines in catalytic applications has been limited by their poor solubility in water.57 The ability to produce heterogeneous catalysts with poorly water soluble metal macrocycles such as heme b and metallophthalocycanines provides an avenue to accessing their catalytic properties within aqueous systems. We will investigate the properties of these materials in future work. Conclusions In this work we demonstrate the engineering of a solid-state metalloprotein material that can be precisely tuned for a variety of different functions at four different levels; namely the metal center, the organic macrocycle, the protein scaffold and the material structure (Figure 6). These solid-state metalloproteins are remarkably stable and reusable. The ability to produce large quantities, the ease of manipulating the protein sequence, and the long term stability of materials made from this protein make recombinant honeybee silk a promising template for de novo engineering of a wide range of biomimetic metalloprotein materials with diverse functional properties. Supporting Information Mutagenesis primers, FTIR spectra of silk films with and without heme-b, AmelF3 sequences and additional data supplied as Supporting Information. We have also supplied a video demonstrating the preparation of heme-silk sponges. This material is available free of charge via the Internet at http://pubs.acs.org. Acknowledgements We wish to thank Dr Gunjan Pandey for his assistance in photographing the silk sponges, Ms Andrea Woodhead for her assistance in collecting the Raman spectra, Dr Alagacone 17 ACS Paragon Plus Environment


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Sriskantha for preparing honeybee silk protein samples, Dr Peter Campbell for his assistance in the preparation of this manuscript and Dr Andrew Warden for discussions prior to publication. Samples of silkworm silk were a gift from Prof David Kaplan. Author information These authors contributed equally to this work. Trevor D. Rapson & Tara D. Sutherland Corresponding authors *

[email protected]

*

[email protected]

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For Table of contents use only De novo engineering of solid-state metalloproteins using recombinant coiled-coil silk Trevor D. Rapson, Tara D. Sutherland, Jeffrey S. Church, Holly E. Trueman, Helen Dacres and Stephen C. Trowell


De Novo Engineering of Solid-State Metalloproteins Using

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